Isotope tracing reveals bacterial catabolism of host-derived glutathione during Helicobacter pylori infection

Mammalian cells synthesize the antioxidant glutathione (GSH) to shield cellular biomolecules from oxidative damage. Certain bacteria, including the gastric pathogen Helicobacter pylori, can perturb host GSH homeostasis. H. pylori infection significantly decreases GSH levels in host tissues, which has been attributed to the accumulation of reactive oxygen species in infected cells. However, the precise mechanism of H. pylori-induced GSH depletion remains unknown, and tools for studying this process during infection are limited. We developed an isotope-tracing approach to quantitatively monitor host-derived GSH in H. pylori-infected cells by mass spectrometry. Using this method, we determined that H. pylori catabolizes reduced GSH from gastric cells using γ-glutamyl transpeptidase (gGT), an enzyme that hydrolyzes GSH to glutamate and cysteinylglycine (Cys-Gly). gGT is an established virulence factor with immunomodulatory properties that is required for H. pylori colonization in vivo. We found that H. pylori internalizes Cys-Gly in a gGT-dependent manner and that Cys-Gly production during H. pylori infection is coupled to the depletion of intracellular GSH from infected cells. Consistent with bacterial catabolism of host GSH, levels of oxidized GSH did not increase during H. pylori infection, and exogenous antioxidants were unable to restore the GSH content of infected cells. Altogether, our results indicate that H. pylori-induced GSH depletion proceeds via an oxidation-independent mechanism driven by the bacterial enzyme gGT, which fortifies bacterial acquisition of nutrients from the host. Additionally, our work establishes a method for tracking the metabolic fate of host-derived GSH during infection.


Introduction
Oxidative stress results from an imbalance between the production and detoxification of reactive oxygen species (ROS) in a biological system. ROS production is a hallmark of microbial infections, with epithelial barriers, phagocytic immune cells, and pathogen-associated virulence factors all contributing to oxidative stress at the host-microbe interface. ROS accumulation in infected tissues can induce the oxidation of important cellular components including proteins, lipids, and DNA [1,2]. Host cells shield these biomolecules from oxidative damage in part by synthesizing the antioxidant glutathione (GSH) [2]. As the most abundant molecular thiol in eukaryotes, GSH is essential for the detoxification of ROS, the reduction of protein thiols, and the maintenance of intracellular redox homeostasis [3]. Consequently, GSH deficiency has been associated with severe pathophysiological outcomes including neurodegenerative diseases [4][5][6] and metabolic disorders [7][8][9].
Microbial utilization of host GSH during infection can also contribute to disease. The facultative intracellular pathogen Francisella tularensis metabolizes host GSH to support its replication within mammalian cells [10]. Listeria monocytogenes uses host GSH as an environmental cue to regulate virulence gene expression [11,12]. Several extracellular pathogens, including Streptococcus mutans [13] and Campylobacter jejuni [14,15], consume exogenous GSH for nutritional purposes. While genetic approaches have been valuable for identifying bacterial genes required for GSH import and metabolism, tools for directly assessing bacterial utilization of host GSH remain limited. To date, most studies have relied on metabolic labeling of bacterial cultures with radiolabeled GSH to measure GSH uptake [16]. The ability to directly monitor bacterial metabolism of host-derived GSH in the context of infection would provide new mechanistic insights into microbial utilization of this important cellular thiol.
The gastric pathogen Helicobacter pylori, a major cause of peptic ulcers and the leading risk factor for gastric cancer [17][18][19], offers a powerful model system for studying GSH dynamics at the host-microbe interface. H. pylori infection significantly decreases GSH levels in human gastric cells and tissues [20][21][22][23][24][25], which has been attributed to the persistent generation of ROS during infection [20,23,25]. H. pylori induces ROS production by recruiting ROS-generating immune cells to infected tissues, stimulating the activity of host oxidases, and secreting virulence factors that increase ROS levels within infected cells [20,26,27]. Oxidative stress plays a significant role in the development of H. pylori-induced gastric pathology and cancer; indeed, the genetic disruption of host oxidases [28], or the oral administration of antioxidants [29], significantly reduces gastric inflammation [29], colonization [30], and/or tumor formation [31] in rodent models of infection. While the oxidizing conditions that accompany H. pylori infection have been hypothesized to lower host GSH levels by converting GSH to its oxidized form [20], the precise mechanism underlying host GSH depletion remains unresolved.
In this study, we combined isotope tracing with reactivity-guided metabolomics to quantify the abundance and metabolism of host-derived GSH during H. pylori infection. We found that the H. pylori enzyme γ-glutamyl transpeptidase (gGT), which hydrolyzes GSH to glutamate and cysteinylglycine (Cys-Gly) [16,32], is required for the depletion of intracellular GSH from H. pylori-infected cells. Our results indicate that GSH depletion is driven by bacterial catabolism, and not by oxidation, of this abundant host thiol during H. pylori infection.

H. pylori depletes GSH from gastric cells
We measured the concentration of reduced GSH in three gastric cancer cell lines (AGS, MKN28, and NCI-N87) infected with H. pylori strain G27. Consistent with prior studies [20][21][22][23][24][25], we observed a significant decrease in the intracellular GSH concentration of H. pylori-infected cells relative to uninfected controls (Fig 1A-1C). Both infected and uninfected cells were equally viable under the tested conditions (multiplicity of infection (MOI) 50 for 10 h), demonstrating that GSH depletion during H. pylori infection was not due to an overall decrease in gastric cell viability. GSH levels decreased over time in H. pylori-infected AGS cells, with significantly lower concentrations evident as early as 7 h post infection ( Fig 1D). These findings indicate that H. pylori infection induces a progressive decline in host GSH levels, independent of host cell death.

H. pylori does not increase GSH oxidation in gastric cells
GSH depletion during H. pylori infection has previously been attributed to the accumulation of ROS within infected cells [20,23,25]. H. pylori infection is known to stimulate ROS production [25,[33][34][35], which could in principle deplete intracellular pools of reduced GSH by generating the oxidized disulfide dimer GSSG. Indeed, H. pylori-infected AGS cells exhibit substantially higher levels of intracellular ROS than uninfected controls [25,33,34]. To determine whether oxidation is the primary cause of GSH depletion during H. pylori infection, we quantified total GSH (reduced GSH plus oxidized GSSG) levels in H. pylori-infected AGS cells. Similar to reduced GSH (Fig 1A-1C), we observed a significant decrease in total GSH following H. pylori infection (Fig 2A). Notably, GSSG levels did not increase in H. pylori-infected cells, suggesting that the observed depletion of total GSH was not due to GSH oxidation and instead driven by a specific decrease in reduced GSH. To validate our findings, we measured the total GSH concentration of H. pylori-infected cells treated with the potent reducing agents 2-mercaptoethanol or N-acetylcysteine. Treatment with 2-mercaptoethanol increased the overall GSH concentration of uninfected cells (Fig 2B, compare bars 1 and 3; S1 Fig), as previously described [36]; however, neither reducing agent inhibited the loss of GSH induced by H. pylori infection (Fig 2B). Taken together, these results support that H. pylori infection depletes host GSH via an oxidation-independent mechanism.

H. pylori gGT is required for host GSH depletion
As an extracellular gastric pathogen, H. pylori has access to dietary nutrients in the stomach and relies on the uptake of host amino acids for survival [37][38][39][40]. The import of extracellular glutamine is facilitated by its deamidation to glutamate, a process catalyzed by the periplasmic enzyme gGT [41]. gGT also generates glutamate through the hydrolysis of γ-glutamyl compounds including GSH, which is converted to glutamate and Cys-Gly [16,32]. Although gGT is known to hydrolyze extracellular GSH, it is unknown whether this enzyme contributes to GSH depletion within gastric cells infected by H. pylori. To test this, we compared the amount of reduced GSH in AGS cells infected with wild-type (WT) H. pylori G27, an isogenic gGT deletion mutant (ΔgGT), or a genetically complemented mutant strain (ΔgGT::gGT). Consistent with our prior findings (Fig 1A-1C), we detected significantly lower levels of reduced . GSH data are expressed as a percentage of the average reduced GSH concentration in the corresponding uninfected control. Data represent three independent experiments. Each circle represents an independent experiment. Error bars represent means ± SD. **P < 0.01; ***P < 0.001; ns, not significant. Two-tailed unpaired t-tests were used for (A-C) and multiple unpaired t-tests were used for (D).  . AGS cells were also incubated with BME, NAC, or medium alone for the duration of infection. Data represent three independent experiments. Each circle represents an independent experiment. Error bars represent means ± SD. **P < 0.01; ***P < 0.001; ****P < 0.0001; ns, not significant. A two-tailed unpaired t-test was used for (A), and a two-way ANOVA with Šídák's multiple comparisons test was used for (B).  (Fig 3A) to the ΔgGT mutant. Notably, infection with H. pylori mutants lacking the secreted virulence factors cytotoxin-associated gene A (ΔcagA) or vacuolating cytotoxin A (ΔvacA), which have previously been shown to stimulate ROS production in infected cells [25,35,42], diminished host GSH by the same extent as WT H. pylori. These findings underscore that GSH depletion is not driven by oxidation under the tested conditions (Fig 2). In addition, because the WT, ΔgGT, and ΔgGT::gGT strains exhibited uniform growth kinetics when cultured individually (Fig 3B), and all strains were similarly viable when co-cultured with AGS cells (S3 Fig), the observed changes in host GSH levels were not due to represents an independent experiment. Error bars represent means ± SD. *P < 0.05; ***P < 0.001; ****P < 0.0001; ns, not significant. A one-way ANOVA with Šídák's multiple comparisons test was used for (A) and (C), and a oneway ANOVA with Dunnett's multiple comparisons test was used for (E). https://doi.org/10.1371/journal.ppat.1011526.g003

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Isotope tracing reveals catabolism of host-derived glutathione by Helicobacter pylori differences in the viability of the tested strains. We detected similar changes in the abundance of host GSH using AGS cells infected with a second WT strain of H. pylori, PMSS1, and an isogenic gGT deletion mutant ( Fig 3C). Collectively, these data demonstrate that gGT contributes to the depletion of intracellular GSH from gastric cells during H. pylori infection.

gGT facilitates H. pylori uptake of Cys-Gly
Although gGT-mediated hydrolysis of extracellular GSH is known to facilitate H. pylori uptake of glutamate [16], it is unknown whether H. pylori also utilizes the other product of GSH degradation, Cys-Gly, as a nutrient source. Notably, GSH degradation is a mechanism employed by the intracellular pathogen F. tularensis to scavenge Cys-Gly from the host cytosol [43]. Similarly, we hypothesized that H. pylori gGT may mediate bacterial acquisition of Cys-Gly from extracellular GSH. To test this, we cultured WT, ΔgGT, and ΔgGT::gGT H. pylori in rich medium supplemented with isotopically labeled [ 13 C 2 , 15 N 1 ]-GSH and quantified the relative abundance of [ 13 C 2 , 15 N 1 ]-Cys-Gly in bacterial extracts by liquid chromatography-mass spectrometry (LC-MS). All three strains grew equally well following supplementation with isotopically light GSH or [ 13 C 2 , 15 N 1 ]-GSH (S4 and S5 Figs), suggesting that neither cysteine, glycine, nor glutamate were limiting under the tested conditions. Bacterial cell extracts were treated with the thiol-alkylating agent monobromobimane (mBBr) to facilitate the detection of Cys-Gly and other thiol-containing metabolites. The bimane derivative of [ 13 C 2 , 15 N 1 ]-Cys-Gly was readily detected in WT and ΔgGT::gGT H. pylori extracts, but was present at much lower levels in extracts of the ΔgGT mutant (Fig 3D and 3E). These results demonstrate that H. pylori gGT facilitates bacterial acquisition of Cys-Gly from extracellular GSH and suggest that the depletion of host GSH by H. pylori may enable bacterial consumption of Cys-Gly during infection.

H. pylori metabolizes GSH produced by gastric cells
We developed an isotope-tracing assay to determine whether host GSH is hydrolyzed by gGT during H. pylori infection ( Fig 4A). Previous studies have shown that cells supplemented with isotopically heavy glycine can use native biosynthetic pathways (S6 Fig) to generate heavy GSH [44][45][46]. By applying LC-MS to quantify the relative abundance of heavy GSH and heavy Cys-Gly in H. pylori-infected cells, we reasoned we could directly measure bacterial metabolism of host GSH during infection.
To assess bacterial degradation of heavy GSH, we pretreated AGS cells with 50 μg/mL [2-13 C 1 ]-Gly for 24 h, then washed the cells to remove exogenous [2-13 C 1 ]-Gly prior to infection with WT, ΔgGT, or ΔgGT::gGT H. pylori G27 (MOI 35 for 16 h) (Fig 4A). The relative abundance of heavy GSH and heavy Cys-Gly in H. pylori-infected cell extracts was quantified by mBBr labeling and LC-MS analysis. Consistent with our findings using a commercial assay for GSH quantitation (Fig 3A), we detected considerably lower levels of mB-[ 13 C 1 ]-GSH in cells infected with WT or ΔgGT::gGT H. pylori compared to cells infected with the mutant strain (Figs 4D, 4E and S8). Accordingly, we hypothesized that [ 13 C 1 ]-Cys-Gly levels would be greatest in extracts from cells infected with WT or ΔgGT::gGT H. pylori, reflecting gGT-dependent hydrolysis of [ 13 C 1 ]-GSH. Indeed, significant levels of [ 13 C 1 ]-Cys-Gly were only detected in the presence of gGT-encoding H. pylori strains (Fig 4F and 4G). Extracts from cells infected with ΔgGT H. pylori or uninfected controls contained almost no [ 13 C 1 ]-Cys-Gly, suggesting that the production of this metabolite stems from the GSH-degrading activity of adherent represents an independent experiment. Error bars represent means ± SD. *P < 0.05; **P < 0.01; ***P < 0.001; ****P < 0.0001; ns, not significant. A two-tailed unpaired t-test was used for (C), and a two-way ANOVA with Šídák's multiple comparisons test was used for (E) and (G). https://doi.org/10.1371/journal.ppat.1011526.g004

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Isotope tracing reveals catabolism of host-derived glutathione by Helicobacter pylori bacteria. Notably, we were unable to detect [ 13 C 1 ]-Cys-Gly in nonadherent bacteria harvested from the cell culture medium under these conditions (S9 Fig). Altogether, our results demonstrate that gGT degrades host GSH during H. pylori infection and that this process contributes to the depletion of GSH within infected cells.

Discussion
In this study, we found that H. pylori catabolizes host GSH, resulting in the depletion of this protective thiol from H. pylori-infected gastric cells. Although oxidative stress has previously been cited as the presumptive cause of GSH depletion during H. pylori infection [20,23], our findings support a model wherein H. pylori uses the periplasmic enzyme gGT, an established virulence factor, to degrade and acquire nutrients from host GSH. Extensive work by several groups over the past two decades has uncovered multiple immunomodulatory roles for gGT during H. pylori infection, including the inhibition of T-cell proliferation [47] and induction of dendritic cell tolerance [48]. However, to date, no direct link between this enzyme and H. pylori consumption of host GSH has been established.
Our data position gGT at the helm of a nutrient acquisition system that supports bacterial survival in the stomach while corrupting the ability of host cells to maintain intracellular redox homeostasis. Given that GSH is typically maintained at millimolar levels within gastric tissues [49], this tripeptide constitutes a significant reservoir of cysteine and glutamate in the host environment. Indeed, exogenous peptides and amino acids serve as an important carbon source for H. pylori [37,38]. Extracellular glutamate is incorporated into the H. pylori tricarboxylic acid cycle [16], and we found that Cys-Gly, the other product of GSH hydrolysis [16,32], is also internalized by H. pylori (Fig 3E). Although GSH supplementation did not enhance the growth of gGT-expressing strains in broth cultures (S4 Fig), the rich medium required for H. pylori growth could mask nutritional advantages conferred by GSH metabolism in vitro [50]. Nevertheless, given that the expression of gGT is required for H. pylori colonization of murine gastric tissues [48,51], our findings imply that degradation of host GSH supports H. pylori metabolism and survival in vivo.
While our data establish a role for H. pylori gGT in depleting GSH from gastric cells, it remains unclear how H. pylori gains access to intracellular pools of host GSH. The detection of host-derived Cys-Gly in AGS cells infected with WT or ΔgGT::gGT H. pylori, but not in cells infected with the ΔgGT mutant ( Fig 4G) or in WT bacteria harvested from conditioned co-culture medium (S9 Fig), suggests bacterial degradation of host GSH may be contact-dependent. Host cell lysis induced by H. pylori infection could make intracellular GSH available for bacterial consumption. Alternatively, H. pylori may actively translocate GSH from, or stimulate GSH efflux by, infected cells as a means of promoting GSH catabolism by adherent bacteria. Additional studies are needed to determine whether specific H. pylori or host transporters facilitate bacterial access to intracellular GSH.
Although gGT appears to be the principal driver of host GSH depletion during H. pylori infection, other bacterial and/or host factors may augment this phenotype. Indeed, in certain experiments, we detected lower GSH levels in cells infected with the ΔgGT mutant than in uninfected controls (Figs 3C and 4E). Prior studies indicate that the expression of cation transport regulator 1 (CHAC1), a host enzyme that cleaves GSH into 5-oxoproline and Cys-Gly [52], is elevated in H. pylori-infected gastric cells under certain conditions [53,54]. However, there have been conflicting reports regarding the transcriptional regulation of CHAC1 during infection [55], and our data suggest that hydrolysis of host GSH to Cys-Gly in H. pyloriinfected cells is gGT-dependent (Fig 4G). Similarly, the secreted H. pylori virulence factor VacA has been reported to decrease levels of host GSH by increasing the accumulation of intracellular ROS [25], though our data do not support a role for VacA in GSH depletion under the conditions tested here (Fig 3A). Nevertheless, strain-specific factors and infection conditions may influence the mechanisms of GSH depletion that are active at the hostmicrobe interface.
Our work establishes the utility of isotope tracing and mass spectrometry in tracking GSH metabolism during infection. This approach permits the precise monitoring of host-synthesized GSH and its degradation product Cys-Gly in H. pylori-infected cells, thereby offering a powerful complement to conventional radiolabeling assays. While radiolabeled GSH has been extensively used to detect cellular uptake of GSH [56][57][58], radiolabeling approaches can only provide a nonspecific readout of GSH metabolism based on the intracellular accumulation and/or proteome-wide incorporation of a specified radionuclide. These methods are generally unable to resolve the exact metabolites resulting from degradation of a radiolabeled precursor. Furthermore, because it is difficult to assess the distribution of radioactivity in a complex system, radioisotope-based approaches have thus far been limited to studies of GSH uptake by bacteria cultured independently in vitro [10,16,43]. By contrast, isotope tracing coupled to LC-MS enables the targeted analysis of host-derived Cys-Gly in infected cells and provides a safer alternative to radiolabeling approaches. Given the growing list of pathogens that co-opt host GSH to fortify bacterial metabolism [10,[13][14][15]59] or activate virulence mechanisms during infection [12,[60][61][62], this approach should be broadly useful for future studies of GSH utilization by bacteria in the host environment.
In summary, our findings redefine H. pylori-induced depletion of host GSH as a microbedriven metabolic process that facilitates bacterial acquisition of essential amino acids. GSH degradation depletes gastric cells of this protective thiol, sensitizing host tissues to oxidative damage. As such, GSH depletion by H. pylori not only dynamically shapes redox homeostasis at the host-microbe interface, but comprises a previously unappreciated mechanism of 'nutritional virulence' [63] that could potentially be targeted to mitigate H. pylori-associated pathology.

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Isotope tracing reveals catabolism of host-derived glutathione by Helicobacter pylori described previously [64] were a gift from Dr. Osamu Nagano (Division of Gene Regulation, Institute of Advanced Medical Research, Keio University School of Medicine). MKN28 cells were grown in RPMI 1640 medium supplemented with 10% (v/v) FBS-HI (Biowest) and 1% (v/v) ABAM.

Generation of H. pylori mutant strains
H. pylori mutant strains were generated via natural transformation of H. pylori G27 with linear PCR constructs containing the flanking regions of the targeted gene and an antibiotic resistance cassette as previously described [65,66]. To construct H. pylori G27 ΔvacA,~600-base pair (bp) DNA sequences flanking vacA (GenBank HPG27_840) were PCR-amplified from H. To construct H. pylori G27 ΔcagA, a linear deletion construct consisting of the cat resistance cassette flanked by the~500-bp DNA sequences immediately upstream and downstream of cagA (GenBank HPG27_507) was amplified from H. pylori G27MA ΔcagA gDNA using the primers MB-71 and MB-74. The downstream flank of cagA included 43 bp of the 3' end of the cagA ORF. The resulting PCR product was purified using the QIAQuick PCR Purification kit (QIAGEN) prior to transformation of H. pylori G27.
The H. pylori G27 ΔgGT::gGT complemented strain was generated via natural transformation of H. pylori G27 ΔgGT [47] with a linear PCR construct containing the~500-bp DNA sequence immediately upstream of gGT (GenBank HPG27_1063) followed by the complete gGT coding region, the cat cassette, and the~500-bp sequence immediately downstream of gGT. This construct was amplified from plasmid pUC57_gGT::gGT-cat, which was custom synthesized by GenScript, and purified using the QIAQuick Gel Purification Kit (QIAGEN) prior to transformation of H. pylori G27 ΔgGT. All mutant and complemented strains were confirmed by locus-specific PCR reactions and DNA sequencing.
To verify the loss of gGT expression by H. pylori G27 ΔgGT, total RNA was extracted from log-phase cultures (OD 600~0 .4) of WT, ΔgGT, and ΔgGT::gGT H. pylori G27 using the Direct-zol RNA Miniprep Plus kit (Zymo Research) for reverse transcription-quantitative PCR (RT-qPCR) analysis. gGT mRNA was amplified using primers MJB-21 and MJB-22 and the KAPA SYBR FAST One-Step qRT-PCR kit (Kapa Biosystems) according to the manufacturer's instructions. The H. pylori housekeeping gene ppk (polyphosphate kinase, GenBank HP_1010; primers ABS-75 and ABS-76; [67]) was used to normalize mRNA expression. Samples were analyzed using a CFX96 Real-Time PCR Detection System (Bio-Rad). Relative changes in gGT expression were determined using the ΔΔCt method [68].

H. pylori infection of gastric cells
Gastric cancer cell lines were seeded at a density of 2 x 10 5 cells in the appropriate medium (DMEM or RPMI 1640 supplemented with 10% (v/v) FBS-HI) 20-24 h prior to infection in either 24-well flat-bottom tissue culture-treated plates (Corning; luminescence-based GSH assay) or 100-mm tissue culture-treated dishes (Corning; LC-MS analyses). H. pylori overnight

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Isotope tracing reveals catabolism of host-derived glutathione by Helicobacter pylori cultures were either diluted to OD 600 = 0.3 and incubated for an additional 1 h at 37˚C with shaking at 100 rpm under microaerophilic conditions (luminescence-based GSH assay) or centrifuged (18,000 x g, 5 min, room temperature) and resuspended in fresh medium at OD 600 = 1.0 (LC-MS analyses) prior to infection. Bacterial cultures, or an equivalent volume of 10% (v/v) FBS-HI in Brucella broth (mock infection control), were diluted in co-culture medium (DMEM or RPMI 1640 supplemented with 10% (v/v) Brucella broth and 5% (v/v) FBS-HI) to achieve an MOI of 35 or 50. The gastric cell culture medium was replaced with the appropriate co-culture medium, and the infected cells were incubated at 37˚C in a humidified 5% CO 2 incubator for 3-20 h as specified. To verify the MOI and assess bacterial growth during infection, a portion of the H. pylori-containing co-culture medium was collected at the start and end of each experiment and serially diluted in Brucella broth supplemented with 10% (v/v) FBS-HI for the enumeration of bacterial CFU. Gastric cell viability was quantified by Trypan Blue (Thermo Fisher Scientific) exclusion using a Countess II FL automated cell counter (Thermo Fisher Scientific).

Antioxidant treatment
AGS cells were pretreated with either 100 μM or 500 μM 2-mercaptoethanol (Sigma) or 5 mM N-acetylcysteine (Sigma) in DMEM containing 10% (v/v) FBS-HI for 1 h prior to infection. At the time of infection, the cell culture medium was replaced with co-culture medium containing an equivalent concentration of either 2-mercaptoethanol or N-acetylcysteine.

Measurement of intracellular GSH via luminescence-based assay
Gastric cells were washed once with Dulbecco's PBS (DPBS; HyClone) and collected via dissociation with TrypLE Express (Gibco). Cells were pelleted by centrifugation (300 x g, 3 min, room temperature in 15-ml conical tubes (Corning), followed by 21,000 x g, 2 min, room temperature in microcentrifuge tubes). Harvested cells were resuspended in PBS (Gibco), transferred to white-walled 96-well plates (Corning), and analyzed using the GSH-Glo Assay kit (Promega) according to the manufacturer's instructions. Intracellular GSSG and total GSH concentrations were measured using the GSH/GSSG-Glo Assay kit (Promega) according to manufacturer's instructions. Luminescence was measured using a SpectraMax i3x Multi-Mode Microplate Reader.

H. pylori growth curves
Overnight cultures of WT, ΔgGT, and ΔgGT::gGT H. pylori G27 were grown for 16 h at 37˚C and diluted to OD 600 = 0.1 in fresh Brucella broth containing 10% (v/v) FBS HI with or without 50 μg/mL light GSH (Sigma). Subcultures were grown for an additional 22 h at 37˚C. CFU were enumerated directly after subculturing (t = 0) and again following 4, 8, and 22 h of growth.

Metabolic labeling of H. pylori with [ 13 C 2 , 15 N 1 ]-GSH
Overnight cultures of H. pylori were grown for 16 h at 37˚C and diluted to OD 600 = 0.1 in Brucella broth containing 10% (v/v) FBS HI and 50 μg/mL glutathione (glycine-13 C 2 , 15 N) (95% + purity, Cambridge Isotope Laboratories; aka [ 13 C 2 , 15 N 1 ]-GSH). Subcultures were grown for an additional 8 h at 37˚C, and a portion of each culture was collected for the enumeration of CFU. Cultures were then normalized by OD 600 and pelleted by centrifugation (18,000 x g, 10 min, room temperature). Cell pellets were washed twice with PBS (Fisher Scientific) and stored at -80˚C prior to further analysis.

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Isotope tracing reveals catabolism of host-derived glutathione by Helicobacter pylori

Metabolic labeling of gastric cells with [2-13 C 1 ]-Gly
AGS cells were treated with 0-50 μg/mL 2-13 C glycine (99% purity, Cambridge Isotope Laboratories; aka [2-13 C 1 ]-Gly) in DMEM supplemented with 10% (v/v) FBS HI for 24 h. To assess biosynthesis of [ 13 C 1 ]-GSH, AGS cells were washed twice in DPBS (Hyclone), and collected via dissociation with TrypLE Express. Cells were pelleted by centrifugation (300 x g, 3 min, room temperature in 15-ml conical tubes (Corning), followed by 21,000 x g, 2 min, room temperature in microcentrifuge tubes) and stored at -80˚C until further analysis. To assess [ 13 C 1 ]-GSH or [ 13 C 1 ]-Cys-Gly levels in cells co-cultured with H. pylori, AGS cells pretreated with 0 or 50 μg/mL [2-13 C 1 ]-Gly were subsequently washed twice with DPBS and infected at an MOI of 35 for 16 h. Following infection, conditioned cell culture medium containing non-adherent H. pylori was collected and pelleted by centrifugation (18,000 x g, 10 min, room temperature). Bacterial cell pellets were stored at -80˚C until further analysis. AGS cells were treated with 400 μg/mL kanamycin (Thermo Fisher Scientific) in DMEM for 1 h and washed once with DPBS. Cells were collected via dissociation by TrypLE Express and pelleted by centrifugation (300 x g, 3 min, room temperature in 15-ml conical tubes, followed by 21,000 x g, 2 min, room temperature in microcentrifuge tubes) and stored at -80˚C until further analysis.

Metabolite extraction and thiol labeling
Frozen H. pylori or AGS cell pellets were thawed on ice and resuspended in 500 μL ice-cold thiol extraction buffer (50% (v/v) acetonitrile (ACN; Millipore), 50 mM HEPES pH 8 (Sigma), 1 mM TCEP (Sigma), and 1 mM diethylenetriaminepentaacetic acid (DTPA; Sigma)) for 30 min at -20˚C. Samples were centrifuged (21,000 x g, 10 min, 4˚C), and the supernatants were subsequently transferred to glass vials (Fisher Scientific). L-cysteine (1-13 C) (99% purity, Cambridge Isotope Laboratories; 1 μg) was added to each sample and used as an internal standard to confirm similar processing across all replicates. Samples were treated with 35 μL of a 74 mM solution of monobromobimane (mBBr; Santa Cruz Biotechnology) in 100% ACN for 15 min at 60˚C, protected from light, then frozen and lyophilized overnight at -80˚C. Samples were reconstituted in 100% methanol (Avantor) and centrifuged (21,000 x g, 10 min, 4˚C). Supernatants were analyzed immediately thereafter by LC-MS or LC-MS/MS.